SUMMARY

In photosynthesizing plant cells, chloroplasts change their arrangements
and/or positions in response to light irradiation. These photo-orientation
movements of chloroplasts are believed to play important roles in optimizing
the photosynthetic activity of plant cells. We have been investigating the
roles of the actin cytoskeleton in the intracellular movement and positioning
of chloroplasts using the aquatic monocot Vallisneria gigantea
Graebner and the terrestrial dicot Spinacia oleracea L. (spinach). In
Vallisneria epidermal cells, chloroplasts accumulate on the
cytoplasmic layer facing the top surface (outer periclinal layer) under dim
red light, whereas they move to the cytoplasmic layer perpendicular to the
outer periclinal layer (anticlinal layer) under strong blue light. Concomitant
with these responses, actin filaments exhibit dramatic changes in their
configurations. The possible modes of action of the actin cytoskeleton to
regulate the movement and positioning of chloroplasts are briefly summarized,
together with our recent analysis of the association of actin filaments with
chloroplasts isolated from spinach leaves.

Photo-orientation movement of chloroplasts

Chloroplasts, which are cell organelles specifically differentiated for
photosynthesis, change their intracellular arrangements and/or positions in
response to irradiation with light (Senn,
1908; Haupt and Scheuerlein,
1990; Wada and Kagawa,
2001). These movements are widely observed in a variety of plant
species and are designated as the photo-orientation movement of chloroplasts.
Typical examples of such movements are schematically shown in
Fig. 1. Basically, chloroplasts
move to expose their 'faces' to incident dim light and their 'profiles' to
strong light. These movements are believed to play important roles in the
maximization of photosynthetic activity
(Zurzycki, 1955) and the
minimization of photo-damage (Zurzycki,
1957; Park et al.,
1996) in plants under fluctuating light conditions.

Schematic demonstration of photo-orientation movement of chloroplasts in
various types of plant cells. Typical intracellular distributions of
chloroplasts observed under dim or strong light in the coenocytic alga
Vaucheria, the green alga Mougeotia and the angiosperm
Lemna are shown. N, nucleus. Modified from Senn
(1908).

Concomitant with the light-dependent redistribution of chloroplasts, the
spatial reorganization of actin filaments has been shown in the coenocytic
alga Vaucheria (Blatt and Briggs,
1980; Blatt et al.,
1980), the green algae Caulerpa
(Menzel and Elsner-Menzel,
1989) and Mougeotia
(Mineyuki et al., 1995) and
the pteridophytes Selaginella (Cox
et al., 1987) and Adiantum
(Kadota and Wada, 1992a).
Chloroplast movement in the green alga Dichotomosiphon is
microtubule-dependent (Maekawa and Nagai
1988); however, chloroplasts that accumulated in cell apices after
photo-orientation movement were found to be associated with numerous fine
bundles of actin filaments (Fig.
2). Also, in vascular plants, actin filaments surrounding
chloroplasts have frequently been observed by light microscopy
(Kobayashi et al., 1987;
Kadota and Wada, 1992a;
Dong et al., 1996;
Kandasamy and Meagher, 1999).
In A. thaliana, disruption of the actin filaments by the anti-actin
drug latrunculin B led to the disruption of the intracellular arrangement of
chloroplasts (Kandasamy and Meagher,
1999). These studies have pointed out the possibility that actin
filaments not only provide tracks for the movement of chloroplasts but also
function to anchor the chloroplasts at proper intracellular positions. The
different roles of the actin filaments may have resulted from the different
signalling pathways from the photoreceptor systems functioning under different
light conditions. We are investigating these aspects in higher plants, using
the monocotyledonous aquatic plant Vallisneria gigantea Graebner and
the dicotyledonous terrestrial plant Spinacia oleracea L.
(spinach).

Accumulation of chloroplasts in the cell apex of the coenocytic green alga
Dichotomosiphon. (A-D) Dim blue light induced the accumulation of
chloroplasts in the apical region of a cylindrical cell of
Dichotomosiphon over a period of 0-90 min. (E) Microtubules (MT;
green) and (F) actin filaments (AF; red) in the apical region filled with
chloroplasts (blue) were visualized using fluorescence-labelled probes. Scale
bar: 20 μm.

Accumulation response of chloroplasts in Vallisneria

In leaf epidermal cells of Vallisneria, chloroplasts accumulate on
the cytoplasmic layer facing the top surface (outer periclinal layer; P side)
under dim light (accumulation response;
Fig. 3A). We frequently
observed that mitochondria also gather on the P side with the accumulated
chloroplasts (Fig. 4),
suggesting mutual metabolic interactions
(Raghavendra et al., 1994).
Using video microscopy, we analysed the mode of movement of chloroplasts
during the accumulation response semi-quantitatively
(Dong et al., 1995). In the
dark, the chloroplasts slowly migrated between the P side and the cytoplasmic
layers perpendicular to the P side (anticlinal layers; A sides) at a constant
rate. The number of chloroplasts that migrated from the A sides to the P side
was almost the same as that migrating in the opposite direction. Within
several minutes of irradiation with dim red light, the rate of migration of
chloroplasts in both directions increased. After approximately 20 min,
however, the rate of migration of chloroplasts from the P side to the A sides
decreased compared with that in the opposite direction, leading to a gradual
increase in the number of chloroplasts on the P side
(Dong et al., 1995). Thus, red
light appeared to initially enhance the motility of each chloroplast and to
later suppress the motility of chloroplasts on the P side but not on the A
sides (Fig. 5).

Accumulation of mitochondria with chloroplasts in Vallisneria
epidermal cells. Paradermal sections of Vallisneria epidermal cells
under dim light, in which mitochondria (Mit) accumulated along the outer
periclinal wall together with chloroplasts (Chl).

Light-dependent changes in motility of chloroplasts in Vallisneria
epidermal cells. The motility of individual chloroplasts was determined after
digitization of images obtained by video microscopy of Vallisneria
epidermal cells under dim red light in the presence (filled symbols) or
absence (open symbols) of dichlorophenyl dimethylurea (DCMU), an inhibitor of
photosynthetic electron transport. Modified from Dong et al.
(1996).

Both far-red light and inhibitors of photosynthesis [dichlorophenyl
dimethylurea (DCMU), atrazine and tetraphenyl boron] antagonized the
red-light-induced accumulation of chloroplasts on the P side. However, the
modes of inhibition were completely different. Far-red light rapidly
suppressed the initial red-light-induced increase in the motility of
chloroplasts. The rates of migration of chloroplasts between the P side and
the A sides promptly returned to the dark control level. By contrast, in the
presence of DCMU, there was hardly any decline in chloroplast motility after
the initial increased motility of chloroplasts by red light
(Fig. 5). In fact, the
increased migration of chloroplasts between the P side and the A sides
continued for a long time and did not decrease as it did after dim-red-light
irradiation in the absence of DCMU. Thus, in either case, the number of
chloroplasts on the P side did not change because no imbalance in the rates of
migration of chloroplasts between the P side and the A sides occurred. We
consequently succeeded in distinguishing the effects of dim red light on the
motility of chloroplasts. Firstly, there is the rapid, red-light and
far-red-light reversible effect. Red light accelerates the motility of
chloroplasts, whereas far-red light inhibits this increased motility. This
effect is thought to be mediated by photoreceptor phytochromes. The other
effect is a much slower, photosynthesis-dependent suppression of the motility
of chloroplasts. We clarified that the reorganization of actin filaments is
involved in the latter photosynthesis-dependent process. Although we
identified a Ca2+-sensitive motor protein activity that interacts
with actin filaments in Vallisneria leaves
(Takagi 1997), its
intracellular localization has not been determined yet. The possible
involvement of the motor protein activity in the regulation of the motility of
chloroplasts remains to be investigated.

Actin-dependent anchoring of chloroplasts

In Vallisneria epidermal cells after incubation in the dark, actin
filaments on the P side formed a loose network
(Dong et al., 1996) composed of
thin bundles (Yamaguchi and Nagai,
1981). Concomitant with the dim-red-light-induced decrease in
motility of chloroplasts on the P side, the configuration of the actin
filaments markedly changed to a honeycomb array, as if the filaments
surrounded each chloroplast (Fig.
6A). Importantly, DCMU suppressed the red-light-induced
reorganization of actin filaments, as well as the decrease in motility of
chloroplasts (Fig. 5). The
chloroplasts themselves seemed to produce signals to regulate the
configuration of actin filaments so that they could settle into the proper
position under dim red light.

After accumulation on the P side under dim red light, the chloroplasts
became considerably resistant to centrifugal force
(Takagi et al., 1991;
Dong et al., 1998). This effect
was antagonized by treatment with cytochalasin, which simultaneously brought
about the complete fragmentation of the honeycomb array of the actin filaments
surrounding the chloroplasts (Dong et al.,
1998). Therefore, in Vallisneria, we demonstrated for the
first time that actin filaments not only drive the movement of chloroplasts
but also anchor the chloroplasts after the photo-orientation movement.

Avoidance response of chloroplasts in Vallisneria

Under strong light, chloroplasts in Vallisneria epidermal cells
accumulate on the A sides (avoidance response;
Fig. 3B). As seen in other
plant species, blue light specifically induces the avoidance response of
chloroplasts (Izutani et al.,
1990). Under video microscopy, chloroplasts on the P side, which
had been apparently motionless, began to sway randomly within a few minutes of
blue light irradiation. Then, the chloroplasts commenced to move more
directionally and migrated towards the A sides. The chloroplasts that moved to
the A sides did not return to the P side; thus, the number of chloroplasts on
the P side rapidly decreased (Fig.
3B). Upon blue light irradiation, the configuration of actin
filaments on the P side changed, this time from a loose network to a more
stretched network composed of thick bundles
(Fig. 6B).

Using microbeam irradiation, we found that the avoidance response of
chloroplasts was induced locally only in the region exposed to blue light.
Chloroplasts in the non-irradiated regions did not change their positions at
all. The reorganization of the actin filaments was also induced only in the
irradiated region, producing a 'hybrid' cell possessing both the actin
filaments of a honeycomb array surrounding the motionless chloroplasts and the
thick, straight bundles that did not come in contact with any chloroplasts
(Fig. 6C). The thick, straight
bundles of actin filaments most probably function as tracks for the
unidirectional migration of chloroplasts from the irradiated region. The
regulation of the configuration of actin filaments by blue light
photoreceptors is under strict spatial control in individual epidermal
cells.

Interaction of chloroplasts with actin filaments in spinach

We used spinach to investigate the possible interaction of chloroplasts
with actin filaments in vitro because the isolation procedures of
chloroplasts from spinach are well established. Moreover, the light-dependent
redistribution of chloroplasts was suggested, based on measurements of
light-induced absorbance changes of the spinach leaves
(Inoue and Shibata, 1974). By
video microscopy, we confirmed that blue light induced the directional
migration of chloroplasts to avoid the incident light in mesophyll cells. Such
movement was reversibly inhibited by cytochalasin, indicating the involvement
of actin filaments. When the mesophyll cells were partially irradiated with
strong blue light, the avoidance response of chloroplasts was induced only in
the irradiated area (Fig. 7).
Blue light often induced the formation of one or two thick cytoplasmic strands
in each irradiated cell. These cytoplasmic strands moved freely from the
non-irradiated area to the irradiated area and vice versa
(Fig. 7). Although the nature
of the light-induced formation of the cytoplasmic strands has not yet been
characterized in detail, in this case, signals from the photoreceptors seem to
be transmitted from the irradiated area to the surrounding non-irradiated
areas.

Because we succeeded in visualizing the actin filaments associated with
chloroplasts in spinach mesophyll cells under dim light, we attempted to
isolate such chloroplasts from the cells. In the cytoplasm obtained by
squeezing manually dissected cells, we occasionally observed chloroplasts
associated with actin filaments (Fig.
8). This suggested a possible direct interaction of chloroplasts
with actin filaments; however, when chloroplasts were isolated after
homogenization of the leaves and Percoll centrifugation, we could not detect
any actin by immunoblotting of the final fraction, which was rich in intact
chloroplasts (Fig. 9; lane 1).
Actin filaments might have been detached from the chloroplasts during the
isolation procedure. Finally, using such isolated intact chloroplasts, which
are apparently free of actin filaments, we examined the possible binding of
the exogenously added skeletal F-actin. As expected, F-actin co-sedimented
with the intact chloroplasts, depending on the incubation time
(Fig. 9; lanes 2 and 3).

Immunoblot analysis of actin associated with isolated chloroplasts from
Spinacia oleracea. The association of actin with the isolated intact
chloroplasts from spinach leaves was examined by immunoblotting of the
chloroplasts immediately after isolation (lane 1), and incubation for 0 min
(lane 2) and 30 min (lane 3) with exogenously added chicken skeletal muscle
F-actin. The arrow indicates the position of the actin band. Molecular markers
are indicated on the left.

Concluding remarks

The entire process involved in the photo-orientation movement of
chloroplasts can be separated into photoperception, signal transduction,
relocation of chloroplasts and anchoring of chloroplasts. The recent
identification of blue-light photoreceptors for chloroplast relocation by
mutant analysis in A. thaliana
(Kagawa et al., 2001;
Jarillo et al., 2001;
Sakai et al., 2001) is
undoubtedly a great breakthrough. On the other hand, by observing cytoskeletal
organization, we surmise that actin filaments may play important roles in the
relocation and anchoring of chloroplasts. From our study, we found that actin
filaments can at least bind to chloroplasts in vitro. However, most
parts of the signal transduction process, particularly the molecular basis for
the regulation of the dynamic behaviour of actin filaments in plant cells
(Staiger, 2000), remain
unsolved.

Furthermore, as has already been pointed out by several eco-physiologists
(Evans and von Caemmerer,
1996), chloroplasts are almost always positioned along the cell
walls facing the intercellular spaces in the leaves
(Fig. 10). Although this may
be interpreted as indicating that chloroplasts have to capture much
CO2 even under spatially limited conditions
(Terashima et al., 1995), we
know nothing about the mechanism of this simple phenomenon. To obtain a much
deeper insight into the photo-orientation movement of chloroplasts- one of the
most precisely regulated cellular responses in plants to environmental
stimuli- much more extensive collaboration among cell biologists and
eco-physiologists is essential.

Intracellular arrangement of chloroplasts in terrestrial plant leaves.
Intracellular arrangement of chloroplasts is demonstrated in a cross-section
of the adaxial part of a Spinacia oleracea leaf (A) and in a
paradermal section of the parenchyma cells of Maesa japonica (B).
Arrowhead in A indicates the stoma, while the yellow circles in B indicate
intercellular spaces. Scale bars: 25 μm.

ACKNOWLEDGEMENTS

I am grateful to all my colleagues who collaborated with me, especially to
the following who provided excellent results for this article: Ichiro
Terashima, Xiajing Dong, Keiko Sugimoto, Nami Sakurai, Kikuko Domoto, Tomoaki
Kumatani and the late Yasuhiko Izutani. Our study was partly supported by
Grants-in-Aid for Scientific Research from the Ministry of Education, Culture,
Sports, Science and Technology, Japan.